Molecular Biology

Transfecting neurons remains technically challenging due to their sensitivity. Conventional methods, such as Lipofectamine 2000 or Lipofectamine RNAiMAX, often result in significant cytotoxicity, which limits their utility. Although lentiviral transfection offers high efficiency, it is hindered by high costs and complex procedures. This experiment employs a small interfering RNA (siRNA)-specific transfection reagent from the Kermey company. This reagent is a novel nanoparticle-based lipid material designed for the efficient delivery of oligonucleotides, including siRNA, into a wide range of cell types. Its efficacy in achieving high transfection efficiency in neurons, however, has not yet been established. After several days of in vitro neuronal culture, researchers can perform a simple transfection procedure using this reagent to achieve robust transfection efficiency. Notably, the protocol does not require medium replacement 6–8 h post-transfection, streamlining the workflow and minimizing cellular stress.

Although protein–protein interactions (PPIs) are central to nearly all biological processes, identifying and engineering high-affinity intracellular binders remains a significant challenge due to the complexity of the cellular environment and the folding constraints of proteins. Here, we present a two-stage complementary platform that combines magnetic-activated cell sorting (MACS)-based yeast surface display with functional ligand-binding identification by twin-arginine translocation (Tat)-based recognition of associating proteins (FLI-TRAP), a bacterial genetic selection system for efficient screening, validation, and optimization of PPIs. In the first stage, MACS-based yeast display enables the rapid high-throughput identification of candidate binders for a target antigen from a large synthetic-yeast display library through extracellular interaction screening. In the second stage, an antigen-focused library is subcloned into the FLI-TRAP system, which exploits the hitchhiker export process of the Escherichia coli Tat pathway to evaluate binder–antigen binding in the cytoplasm. This stage is achieved by co-expressing a Tat signal peptide–tagged protein of interest with a β-lactamase-tagged antigen target, such that only binder–antigen pairs with sufficient affinity are co-translocated into the periplasm, thus rendering the bacterium β-lactam antibiotic resistant. Because Tat-dependent export requires fully folded and soluble proteins, FLI-TRAP further serves as a stringent in vivo filter for intracellular compatibility, folding, and stability. Therefore, this approach provides a powerful and cost-effective pipeline for discovering and engineering intracellular protein binders with high affinity, specificity, and functional expression in bacterial systems. This workflow holds promise for several applications, including synthetic biology and screening of theragnostic proteins and PPI inhibitors.

It is common practice for laboratories to discard clotted blood or freeze it for future DNA extraction after extracting serum from a serum-separating tube. If freezing for DNA extraction, the blood clot is not usually cryopreserved, which leads to cell membrane fragility. In this protocol, we describe steps to isolate high-quality nuclei from leukocytes derived from whole blood samples frozen without a cryoprotective medium. Nuclei isolated from this protocol were able to undergo ATAC (assay for transposase-accessible chromatin) sequencing to obtain chromatin accessibility data. We successfully characterized and isolated B cells and T cells from leukocytes isolated from previously frozen blood clot using Miltenyi’s gentleMACS Octo Dissociator coupled with flow sorting. Nuclei showed round, intact nuclear envelopes suitable for downstream applications, including bulk sequencing of nuclei or single-cell nuclei sequencing. We validated this protocol by performing bulk ATAC-seq.

Reduced representation sequencing (RRS), particularly through restriction site-associated DNA sequencing (RAD-seq), has been widely adopted for whole-genome genotyping due to its cost-effectiveness and cross-species applicability. Nevertheless, conventional RAD-seq approaches are constrained by intricate workflows and substantial labor intensity. These methods predominantly adhere to a “fragment selection precedes library construction” paradigm, wherein DNA fragments adjacent to restriction enzyme cleavage sites are specifically targeted. In contrast, we present an innovative strategy termed inverse restriction site–associated DNA sequencing (iRAD-seq), which implements a reversed workflow, “library construction precedes fragment selection,” to enable efficient enrichment of DNA fragments not associated with restriction sites for genome-wide genotyping. This approach harnesses Tn5 transposase to concurrently fragment genomic DNA and ligate sequencing adapters, followed by pooled processing of hundreds of libraries under a unified batch restriction digestion step. The iRAD-seq workflow thereby achieves significant simplification and enhances operational efficiency in RAD-seq library preparation.

Labeling cells with reporter genes allows researchers to visually identify specific cells and observe how they interact with each other in dynamic biological systems. Even though various labeling methods are now available, a specific description of gene knock-in labeling methods for human trophoblast stem cells (hTSCs) has not been reported. Here, we present a streamlined protocol for labeling hTSCs with the green fluorescent protein (GFP) reporter gene via CRISPR/Cas9-mediated knock-in of the gene into the adeno-associated virus site 1 (AAVS1) safe harbor locus. A commonly used hTSC cell line, CT29, was transfected with a dual plasmid system encoding the Cas9 endonuclease and an AAVS1-targeted guide RNA in one plasmid and a donor plasmid encoding a puromycin resistance gene and GFP reporter gene flanked by AAVS1 homology arms. Puromycin-resistant clonal cells were isolated, and AAVS1 integration was confirmed via PCR and sequencing of the PCR products. The labeled cells are proliferative and can give rise to extravillous cytotrophoblast cells (EVT) and the syncytiotrophoblast (ST). To our knowledge, this is the first report using the CRISPR/Cas9 system for AAVS1 integration of a reporter gene in human trophoblast stem cells. It provides an efficient tool to facilitate the study of human trophoblast development and function in co-culture systems and will be highly useful in developing clinical gene therapy-related plasmid constructs.

Since its introduction, the CRISPR/Cas9 system has been used in many organisms for precise and rapid genome editing, as well as for editing multiple genes at once. This targeted mutagenesis makes it easy to analyze the function of a gene of interest (goi). The standard method for genetic manipulation of the model organism Neurospora crassa has been homologous recombination. It is well established and widely used to create knock-out or overexpression mutants. The recently developed CRISPR/Cas9 system is an addition to the toolkit for genetically manipulating N. crassa. For this protocol, a strain stably expressing the Cas9 endonuclease is required. After designing the gRNA with the online tool CHOP-CHOP, a synthetic gRNA is used to transform macroconidia via electroporation. Combining the goi-gRNA with a gRNA targeting the csr-1 gene as a selection marker allows for easy identification of colonies with mutations at the target site of the goi, since the obtained resistance to Cyclosporin A (CsA) allows for selecting editing events. The mutation type can be detected by PCR of the edited gene region followed by Sanger sequencing. This system is fast and easy to handle, offering an attractive alternative to homologous recombination, especially for targeting multiple genes simultaneously.

Single-cell and single-nucleus RNA sequencing are revolutionizing our understanding of cellular biology. The identification of molecular markers, single-cell transcriptomic profiling, and differential gene expression at the cellular level has revealed key functional differences between cells within the same tissue. However, tissue dissociation remains challenging for non-model organisms and for tissues with unique biochemical properties. For example, the mosquito fat body, which serves functions analogous to mammalian adipose and liver tissues, consists of trophocytes—large, adipocyte-like cells whose cytoplasm is filled with lipid droplets. Conventional enzymatic dissociation methods are often too harsh for these fragile cells, and their high lipid content can interfere with reagents required for single-cell transcriptomic analysis. Single-nucleus RNA sequencing (snRNA-seq) offers an alternative strategy when intact cells with high-quality RNA cannot be obtained by enzymatic or mechanical dissociation. Here, we present an optimized reproducible methodology for nuclei isolation from the fat body of Anopheles gambiae mosquitoes, enabling high-quality snRNA-seq. Our approach involves tissue fixation and lipid removal, followed by cell lysis and nuclei purification using a sucrose cushion. We validated this protocol on both sugar-fed and blood-fed samples, established quality metrics to remove potential ambient RNA contamination, and demonstrated that snRNA-seq using this method yields high-quality sequencing results.

This protocol presents a modified version of the Filterprep method originally reported in New Biotechnology, adding an optional step to reduce endotoxin levels. Filterprep is a simple, rapid, and cost-effective approach to plasmid DNA purification that couples ethanol precipitation with a single spin-column filtration step, eliminating chaotropic salts and silica binding. The formulations and parameters are fully transparent and do not rely on proprietary buffers, using only standard laboratory reagents and widely available miniprep columns. Under matched conditions, the method recovers high-purity plasmid DNA with yields up to fivefold higher than those obtained with representative commercial midiprep kits. The workflow is readily adoptable in most molecular biology laboratories and, under routine conditions, can be completed in approximately 40 min. The resulting DNA is suitable for molecular cloning, PCR, sequencing, and other downstream biochemical applications. Endotoxin is a lipopolysaccharide (LPS) found in the outer membrane of Gram-negative bacteria and may carry over during plasmid preparation. For experiments requiring lower endotoxin input, an optional modification resuspends the DNA pellet in a Triton X-114 wash buffer before column loading to decrease lipopolysaccharide carryover. The method is modular and extensible, allowing adjustment of precipitation and wash conditions, variation in the number of washes, selection of alternative column formats, and integration of endotoxin-reduction modules without altering the core principle. These features facilitate troubleshooting and quality control, enable scaling from routine batches to larger culture volumes and higher throughput, and allow seamless integration with existing workflows.

The exploration of microbial genomes through next-generation sequencing (NGS) and genome mining has transformed the discovery of natural products, revealing an immense reservoir of previously untapped chemical diversity. Bacteria remain a prolific source of specialized metabolites with potential applications in medicine and biotechnology. Here, we present a protocol to access novel biosynthetic gene clusters (BGCs) that encode natural products from soil bacteria. The protocol uses a combination of Oxford Nanopore Technology (ONT) sequencing, de novo genome assembly, antiSMASH for BGC identification, and transformation-associated recombination (TAR) for cloning the BGCs. We used this protocol to allow the detection of large BGCs at a relatively fast and low-cost DNA sequencing. The protocol can be applied to diverse bacteria, provided that sufficient high-molecular-weight DNA can be obtained for long-read sequencing. Moreover, this protocol enables subsequent cloning of uncharacterized BGCs into a genome engineering-ready vector, illustrating the capabilities of this powerful and cost-effective strategy.

A simple and effective method to identify genetic markers of yield response to nitrogen (N) fertilizer among maize hybrids is urgently needed. In this article, we describe a detailed methodology to identify genetic markers and develop associated assays for the prediction of yield N-response in maize. We first outline an in silico workflow to identify high-priority single-nucleotide polymorphism (SNP) markers from genome-wide association studies (GWAS). We then describe a detailed methodology to develop cleaved amplified polymorphic sequences (CAPS) and derived CAPS (dCAPS)-based assays to quickly and effectively test genetic marker subsets. This protocol is expected to provide a robust approach to determine N-response type among maize germplasm, including elite commercial varieties, allowing more appropriate on-farm N application rates, minimizing N fertilizer waste.